Download Cell Culture Training Form

Updated 5/22/12
Tissue Culture Orientation 1. Complete EH&S Biosafety training modules • Biosafety, Bloodborne Pathogens, and Biosafety Cabinets • http://www.colorado.edu/ehs/training/biosafety.html 2. Media supplies storage • Cold room • TC Freezer • Aliquoting 3. TC Room • Supplies (dishes, flasks, PBS, pipettes, tips, conical tubes and epi tubes) • Remember to re-­‐stock 4. Biohazard waste • Sharps -­‐ Sharps (needles, glass Pasteur pipettes, etc) should be disposed of in the plastic sharps containers underneath each hood. • Liquids -­‐ Liquids are aspirated into the vacuum flasks containing bleach. When the flask is full, add more bleach, and pour down the drain. To dispose of media and cell culture vessels outside of the hood, pour bleach into the vessel, swirl, and pour down the drain. • Plastic serological pipettes -­‐ Plastic pipettes are disposed of in the containers under the hoods. • Tips -­‐ Plastic pipette tips need to be double bagged to prevent puncturing the biohazard bag during disposal. Tips are disposed of into the small biohazard bags in the hoods. When the bag is full, it is sealed and placed in the biohazard bag in the pedal containers. • Other -­‐ Anything that does not fit into one of the above categories (e.g. culture dishes, epi tubes) that has been contaminated with biohazardous materials should be placed into the pedal containers. 5. Sterile Technique • Spray 70% EtOH on work area, supplies, and gloves • Arrange items in hood to minimize clutter and airflow obstruction • Do not block air vents – interruption of airflow compromises hood sterility • Only open sterile bottles, tubes, supplies, etc. inside the TC hood • Do not pass anything over open bottles, tubes, plates • Do not touch sterile tips or pipettes to surfaces 6. Cell Culture • Maintenance, passage, and counting Updated 5/22/12
7. CO2 • Tanks are connected to a manifold mounted in the entry way of A391 • Verify that 2 tanks in use + 2 full backup tanks are connected at all times • Tank pressure: full tanks read 800-­‐1000 psi • Line pressure: -­‐ Never change the pressure valve! -­‐ Manifold is set to 15 psi. It is important not to exceed 15 psi or you will damage the incubators. 8. Special notes for undergraduate use of TC • Undergraduates should not aliquot common reagents (e.g. serum, trypsin, pen/strep). It is important that everyone have access to a consistent stock of tissue culture reagents. Graduate students and post-­‐docs will take care to prepare these reagents in a consistent manner. • Undergraduates should not access the cell dewars unless supervised by their mentor Updated 5/22/12
Tissue culture basics Cells must be taken care of properly so they can stay happy and give you good, consistent results! This includes keeping them contamination-­‐free, well fed, and in the proper atmospheric conditions. Typically mammalian cells are kept at 37deg C, 5% CO2 in a humidified incubator, but some cells may be kept at other CO2 levels. The cells we culture are either primary cells (derived directly from the source, undergo senescence within a few passages) or cell lines (cells that have been transformed so they can keep dividing while maintaining the same phenotype). It is best to use cell lines up to the passage recommended, though, since repeated passaging can eventually lead to senescence or other phenotypic changes. Cells are typically cultured on rigid tissue culture dishes or plates (these dishes are treated so the surface is hydrophilic, and cells can adhere to the surface through proteins deposited from the serum). Some cells, however, require coating of dishes with ECM proteins prior to cell seeding. Feeding cells/Media Cells should typically be fed (media exchanged) every 2-­‐3 days if they are in between passages. The media is different for each cell type, but typically they consist of: basal medium (contains essential amino acids, vitamins, glucose, ions, buffers, indicator coloring), serum (isolated from animal plasma and so not well-­‐
defined, but contains nutrients and growth factors required for cell growth), antibiotic (penicillin/streptomycin), and L-­‐glutamine (an essential amino acid that is less stable when stored above freezing than others and so requires supplementation). Media color is often an indication of the culture status: orange/yellow media indicates acid waste product buildup (you should feed the cells before they get to this point!), while pink media indicates the CO2 level is low (check the incubator to make sure the CO2 levels are OK). Passaging cells Cells need to be passaged, or subcultured into a more dilute fraction, every few days so they can continue dividing. The amount of time between passages (and the dilution of each passage) varies between cell types, but in general cells should be passaged before they reach confluence (100% density) to prevent senescence or other changes in cell behavior due to high cell density. 1) Heat media in the water bath, trypsin-­‐EDTA (0.25%) preferably at room temperature (since trypsin can self-­‐digest and lose activity, especially at higher temperatures) for about 20-­‐30min. 2) Aspirate media, rinse cells with PBS (without Ca2+ or Mg2+) to wash out as much media as possible (since serum contains protease inhibitors that can lower the activity of trypsin). 3) Aspirate PBS, add a small amount of trypsin-­‐EDTA so that it just covers the bottom of the dish/flask—this will detach the cells from the dish within 3-­‐5 min at 37degC (this time varies for cell type). For T75 flasks, about 1.5mL is sufficient; for T25s, 0.5mL; for a 6-­‐well plate well, 0.25mL. a. Trypsin is a protease that cleaves proteins involved in cell-­‐matrix and cell-­‐cell bonds. It can be damaging to cells when they are exposed to too much or for too long, so it’s important to use as little trypsin for as short an amount of time as possible. b. EDTA is a metal ion chelator that sequesters Ca2+ required for integrin (cell-­‐matrix) and cadherin (cell-­‐cell) bonds. Updated 5/22/12
4) When the cells are detached (you may need to rap the flask), add the desired amount of media for dilution into subsequent flasks and gently pipet a few times to break up cell clumps and distribute the cells evenly (be sure to avoid bubbles). It is important to add enough media to inactivate the trypsin so it does not damage the cells (a 1:50 dilution is typical). Alternatively, you can suspend the cells in media, centrifuge down (1200rpm, 5min), and resuspend in fresh media, to remove nearly all of the trypsin. Contamination We almost always make our media with antibiotics, so contamination with bacteria is not a common problem. However, if your cultures do become contaminated, you will generally be able to see cloudiness in your media within 18-­‐24h of contamination. If you see contamination, you should dispose of your plate/flask immediately by bleaching it and disposing it in the biohazard. Alert the others in your incubator so they can make sure none of their cultures have become contaminated as well. Alert TC Manager(s) so that a systemic problem can be identified (ex: contaminated stock reagent). Freezing cells To store cells for later use, they can be kept in a solution of culture medium or serum + DMSO (a cryoprotectant) and stored in liquid nitrogen or a -­‐80degC freezer indefinitely. The protocol varies slightly for each cell type, so it’s best to refer to the protocol that usually arrives with a bullet of new cells. Typically, though, cells are trypsinized, resuspended and centrifuged, and resuspended again in 90% serum + 10% DMSO, and vials contain 1mL of cell suspension at about 2-­‐5x106 cells/mL. Place vials in an isopropanol-­‐containing slow-­‐freeze container and store at -­‐80degC overnight, then transfer to liquid nitrogen for long-­‐term storage. Thawing cells 1) Remove from storage in liquid nitrogen or -­‐80degC. Immerse immediately in 37degC water bath (do not immerse cap) for 1-­‐2 min until vial is completely thawed—quick thawing will minimize damage to cell membranes. Wipe vial with ethanol. 2) Slowly pipet vial contents into flask with pre-­‐warmed media, and evenly distribute cells. Change media once cells have attached to remove residual DMSO (toxic to cells). Alternatively, pipet vial contents into tube with media, spin down in centrifuge (1200rpm, 5min), aspirate supernatant to remove DMSO, and resuspend in fresh media. For more detail, refer to Culture of Animal Cells book by R. Ian Freshney, in the lab or available online through the CU library website. Updated 5/22/12
Tissue Culture – Hands-­‐On Training Student: Mentor: This portion of your training is intended to be completed with your graduate student or post-­‐doc mentor. Once completed, please return your signed form to Jen or Melissa. TC Orientation Date: Date: Bloodborne Pathogens Date: Biosafety Cabinets Date: EH&S Biosafety Training Biosafety Practice Exercises with Mentor Observation Exercise #1 – Sterile Technique Practice (to be completed in biosafety cabinet, maintaining sterility throughout) 1. Pipet 30 mls of sterile PBS into a 50ml conical tube. 2. Pipet 10 mls from the 50 ml conical tube into 3 15ml conical tubes. 3. Pipet 15 ul into an epitube. Dispose of the pipet tip. 4. Pipet 150 ul into an epitube. Dispose of the pipet tip. 5. Pipet 500 ul into an epitube. Dispose of the pipet tip. 6. Aspirate PBS from all of the tubes. 7. Dispose of Pasteur pipet, serological pipets, wrappers, and conical tubes. 8. Empty vacuum flask. Date: Student: Mentor: Date: Student: Mentor: Mentor: Exercise #2 – Cell culture and passage 1. Observe your mentor passage cells. Date: Student: 2. Your mentor observes you passage cells. Date: Student: Mentor: Date: Student: Mentor: